aem accepts, published online ahead of print on 30...

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Francisella tularensis subspecies novicida chitinases and Sec secretion system contribute to 10 biofilm formation on chitin 11

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Jeffrey J. Margolis1, Sahar El-Etr2, Lydia-Marie Joubert3, Emily Moore4, Richard Robison4, Amy 15 Rasley2, Alfred M. Spormann5, and Denise M. Monack1* 16

17 Department of Microbiology and Immunology, Stanford University School of Medicine, 18

Stanford, CA 94305, USA1. Bioscience and Biotechnology Division, Lawrence Livermore 19 National Laboratory, Livermore, CA 94550, USA2. Cell Sciences Imaging Facility, Stanford 20

University School of Medicine, Stanford, CA 943053. Department of Microbiology and 21 Molecular Biology, Brigham Young University, Provo, UT 84602, USA4. Department of Civil 22

and Environmental Engineering, Stanford University, Stanford, CA 94304, USA5. 23

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28 Running Title: Francisella biofilm formation on chitin 29

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37 38 39 * Corresponding Author: Mailing address: 299 Campus Dr., Fairchild Bldg. D347, Stanford, CA 40

94305. Phone: (650) 725-1756. Fax: (650) 723-1837. E-mail: [email protected] 41 42

Copyright © 2009, American Society for Microbiology and/or the Listed Authors/Institutions. All Rights Reserved.Appl. Environ. Microbiol. doi:10.1128/AEM.02037-09 AEM Accepts, published online ahead of print on 30 November 2009

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ABSTRACT 1

Francisella tularensis, the zoonotic cause of tularemia, can infect numerous mammals and other 2

eukaryotes. Although studying F. tularensis pathogenesis is essential to comprehending disease, 3

mammalian infection is just one step in the ecology of Francisella species. F. tularensis has 4

been isolated from aquatic environments and arthropod vectors, environments in which chitin 5

could serve as a potential carbon source and as a surface for attachment and growth. We show 6

that F. tularensis subsp. novicida forms biofilms during colonization of chitin surfaces. The 7

ability of F. tularensis to persist using chitin as a sole carbon source is dependent on chitinases, 8

since mutants lacking chiA or chiB are attenuated for chitin colonization and biofilm formation in 9

the absence of exogenous sugar. A genetic screen for biofilm mutants identified the Sec 10

translocon export pathway and 14 secreted proteins. We show that these genes are important for 11

initial attachment during biofilm formation. We generated defined deletion mutants in two 12

chaperone genes (secB1 and secB2) involved in Sec-dependent secretion and in four genes that 13

encode for putative secreted proteins. All mutants were deficient for attachment to polystyrene 14

and chitin surfaces and for biofilm formation compared to wild-type F. novicida. In contrast, 15

mutations in the Sec translocon and secreted factors did not affect virulence. Our data suggest 16

biofilm formation by F. tularensis promotes persistence on chitin surfaces. Further study of the 17

interaction of F. tularensis with the chitin microenvironment may provide insight into 18

environmental survival and transmission mechanisms of this pathogen. 19

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INTRODUCTION 1

Francisella tularensis is a Gram-negative facultative intracellular pathogen that causes 2

the zoonotic disease tularemia (53). Although researchers have focused on various aspects of F. 3

tularensis infections in mammalian hosts, this organism can survive and grow in one of the 4

widest environmental ranges of any studied pathogen. Indeed, F. tularensis has been isolated 5

from a variety of sources including lagomorphs, arthropods, amoeba and fresh water (3, 57, 62, 6

63). Mammals either succumb to infection or clear the bacterium (71) suggesting that mammals 7

may not support prolonged persistence of F. tularensis in nature. Understanding the 8

environmental lifestyle of F. tularensis will help elucidate the survival mechanisms of this 9

pathogen outside of a host and identify risks for human exposure. Recently, outbreaks of 10

tularemia were associated with fresh water, particularly outbreaks of F. tularensis subspecies 11

holarctica (Type B) in Eurasia (11, 81). While the most virulent subspecies, F. tularensis subsp. 12

tularensis (Type A), was historically linked with the arid climates of North America, a recent 13

epidemiological study found that 100% of tularemia mortality was associated with Type A1 14

strains found in moist climates of the United States (40), suggesting that water may serve as an 15

environmental reservoir for F. tularensis. 16

The survival of some bacteria in an aquatic environment is associated with their ability to 17

utilize chitin as a carbon source. Chitin is the second most abundant biopolymer in nature and 18

provides structure to many organisms, including the cell wall of fungi (5) and the exoskeleton of 19

arthropods and insects (51). This oligomer of N-acetyl-D-glucosamine (GlcNAc) is hydrolyzed 20

by a family of enzymes, termed chitinases (6). These enzymes serve a variety of roles and are 21

conserved from bacteria to mammals. Bacterial chitinases provide environmental organisms the 22

ability to acquire carbon in otherwise nutrient-limiting conditions (37). For example, Vibrio 23

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cholerae, the etiological agent of cholera, utilizes chitinases to persist in marine environments on 1

copepod molts (54). The interaction of V. cholerae with chitin influences various metabolic and 2

physiologic responses in this microorganism. For example, Meibom et al. demonstrated that 3

association with chitin and chitin-derivatives led to a specific expression profile in V. cholerae 4

that included two chitinase genes and the pili genes required for colonization and subsequent 5

biofilm formation on nutritive and non-nutritive surfaces (49). Environmental studies have 6

clearly shown that attachment to chitin surfaces is an integral part of the aquatic lifestyle of V. 7

cholerae, and these surface-attached bacterial communities constitute a successful survival 8

mechanism (66). 9

Formation of biofilms is associated with enhanced survival during environmental stress 10

(1) and increased resistance to antibiotics (13). Biofilms formed by many pathogenic bacteria 11

play an important role in environmental persistence and disease transmission. For instance, 12

Yersinia pestis biofilms are reported to function in transmission of plague bacteria via 13

colonization of the proventriculus of fleas and the mouth of nematodes (15, 32). We 14

hypothesized that biofilm formation by F. tularensis may represent a mechanism of persistence 15

and transmission, as well. 16

A review by Hassett et al. (30) indicated that the F. tularensis subsp. holartica live 17

vaccine strain (LVS) can form biofilms on glass coverslips (30). However, the environmental 18

relevance and molecular mechanisms of F. tularensis biofilm formation were not characterized. 19

F. tularensis subspecies encode for 2 conserved putative chitinases, chiA and chiB 20

(http://www.biohealthbase.org). Various F. tularensis subspecies have been isolated from chitin-21

exoskeletoned arthropods (57) and from fresh water, where outbreaks have been associated with 22

chitinous crustaceans (2, 17). We therefore investigated the interaction of F. tularensis subsp. 23

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novicida (F. novicida) with chitin. We show that F. novicida forms biofilms on natural and 1

synthetic chitin surfaces. Formation of these bacterial communities was dependent on two 2

chitinase genes when exogenous sugar was not present. Attachment to chitin was dependent on 3

factors that are secreted by the Sec translocon protein export system. This mechanism of 4

colonization is specific for environmental surfaces, because deletion of genes that facilitate 5

attachment to chitin did not result in defects in virulence. 6

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MATERIALS AND METHODS 1

Bacterial strains and culture conditions. Francisella novicida strain U112 and F. tularensis 2

subsp. holarctica live vaccine strain (LVS), (18, 61), as well as F. novicida Francisella 3

Pathogenecity Island (FPI) and hspX deletion mutants (80), have been previously described. F. 4

tularensis subsp. tularensis strains, SchuS4 and AS2058 (FT-10), were provided by Jean Celli 5

and the New Mexico Department of Health, respectively, and handled under biosafety level-3 6

(BSL-3) precautions per Centers for Disease Control and Prevention protocol. Unless otherwise 7

noted, strains were grown in modified Mueller Hinton media (MMH), (Difco, Corpus Christi, 8

TX) supplemented with 0.025% ferric pyrophosphate, 0.02% IsoVitaleX (Becton Dickinson, 9

Franklin Lakes, NJ) as a cysteine source, and 0.1% glucose as a carbon source. For some 10

experiments, F. tularensis strains were grown in Chamberlain’s Defined Medium (CDM), (47) 11

with or without glucose. For enumeration studies, bacteria were grown on MMH agar plates. 12

Imaging F. novicida colonization on chitin films and sterile crab shell pieces. Wild-type F. 13

novicida was allowed to attach to either synthetic chitin films (82) or sterile crab shell pieces for 14

1 h. After 1 h, surfaces were washed 3X with phosphate buffered saline to remove non-adhered 15

bacteria and samples were incubated at 30°C in CDM without glucose. After one hour and one 16

week of incubation, respectively, crab shell and chitin film samples were processed for scanning 17

electron microscopy (SEM) investigation. Substrates with attached cells were fixed for 3 days at 18

4°C in 4% paraformaldehyde with 2% glutaraldehyde in 0.1M NaCacodylate Buffer (pH 7.3) 19

(EM grade, EMS, Hatfield, PA). After primary fixation, samples were rinsed in the same buffer, 20

post-fixed in 1% aqueous OsO4 for 1 h, and dehydrated in an ascending ethanol series (30, 50, 21

70, 80, 90, 100%; for 20min each), followed by critical point drying with liquid CO2 using a 22

Tousimis SAMDRI-VT-3B apparatus (Tousimis, Rockville, MD). Samples were mounted on 23

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adhesive carbon film on 15mm aluminum stubs, and sputter-coated with 100Å gold/palladium 1

using a Denton Desk 11 TSC Sputter Coater. Visualization was carried out with a Hitachi S-2

3400N VP SEM (Hitachi Ltd, Pleasanton, CA) operated at 10-15kV, working distance 8-10mm, 3

and secondary electron detector. Images were capture in TIF format. 4

Growth in CDM broth. F. novicida was grown overnight in CDM at 37° with aeration. The 5

culture was then diluted to Optical density 600nm (OD600) using an Ultropec 2100 Pro 6

spectrophotomer (Amersham Biosciences, Pittsburgh, PA) 0.05 in either CDM with no sugar, 7

CDM with 10mM glucose or 10mM GlcNAc. Optical density and colony forming units (CFU) 8

were monitored over time for each media condition. The doubling time for each culture was 9

calculated. 10

Imaging of flow cell grown biofilms. Flow cells were assembled as previously described (12, 11

76). The flow system apparatus was sterilized and pre-conditioned with MMH plus 5!g/mL 12

tetracycline (Tet5) overnight at ambient temperature (20-22°C). F. novicida harboring the 13

pKK219-GFP plasmid (26, 41) was grown overnight at 26° C in MMH Tet5 with aeration. 14

Overnight-grown bacteria were diluted 1:50 in fresh media and grown to optical density 600 15

(OD600) 1.0. The culture was then diluted to OD600 0.1. Flow was stopped on the flow system 16

and 1ml of culture was inoculated into each channel of the flow cell. Flow cells were inverted 17

for 1 h to allow the bacteria to adhere. Flow cells were then uprighted and flow was initiated at 18

0.1 ml/minute. Biofilm progression at ambient temperature was imaged by confocal microscopy 19

(Bio-Rad, Hercules, CA) every 24h over the course of 5 days. Z-sections were taken with 0.1 20

µm steps and 3-D renderings of the z-stacks were generated using Volocity imaging software 21

(Improvision, Lexington, MA). 22

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Crystal violet assaying for biofilm formation. Crystal violet assaying for biofilm formation 1

was performed as previously described (59). Briefly, Francisella strains were grown overnight 2

at the appropriate temperature. Cultures were diluted into fresh media to OD600 0.05 and 200!l 3

aliquoted per well in a 96-well polystyrene plate in at least triplicate. The bacteria were allowed 4

to grow statically and sampled at various time points. The OD570 was read in a 96-well 5

microplate reader (BioTek, Winooski, VT). At each time point non-adhered bacteria were 6

removed from the well and 30!l of 0.1% crystal violet was added to each well for 15 minutes. 7

Wells were washed three times with distilled water and the remaining biomass-absorbed crystal 8

violet was solubilized with 95% ethanol. Staining was then quantified at OD570 in a 96-well 9

microplate reader (labeled CV570). All OD readings for the assay comparing relative crystal 10

violet staining between lab strains of Francisella and Type A Francisella were obtained at 11

600nm (CV600) using a NanoDrop spectrophotometer (Thermo Fisher Scientific, Waltham, MA). 12

Transposon library screen for biofilm-deficient mutants. A sequenced two-allele transposon 13

mutant library was used to test for F. novicida transposon mutants that were deficient in biofilm 14

formation (the following reagent was obtained through the NIH Biodefense and Emerging 15

Infections Research Resources Repository, NIAID, NIH: F. tularensis subsp. novicida, “Two-16

Allele” Transposon Mutant Library Plates 1-14, 16-32). The library represents two or more 17

transposon insertions in all non-essential genes. At the time of screening, Plate 15 of the library 18

was unavailable due to quality control issues, resulting in a library size of 2,954 mutants. The 19

two-allele library was received frozen in 96-well format. MMH media was inoculated in 96-well 20

plates with the library and mutants grown overnight to stationary phase at 37°C shaking at 200 21

rpm. Overnight cultures were diluted 1:50 in 200!l of fresh MMH in 96-well plates. Plates 22

were grown statically for 10h in a 37°C incubator and the ability of each transposon-mutant to 23

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form a biofilm was assessed as described above. Mutants exhibiting lower potential for biofilm 1

formation were classified by crystal violet staining more than two standard deviations lower than 2

the plate average. Wild-type F. novicida was included on each plate as a positive control and a 3

well of MMH only was used as a blank. To account for small differences in culture growth, 4

crystal violet staining was normalized to each mutant culture at OD570. Wells where significant 5

growth defects were observed were excluded. Biofilm-deficient transposon-mutants were 6

retested in triplicate. 7

Secondary screening for attachment. Overnight cultures of biofilm mutants identified in our 8

screen were grown in triplicate with shaking (200 rpm) at 37°C. Stationary phase cultures 9

(200!l) were transferred to new 96-well plates and allowed to adhere statically for 1 h at 37°C. 10

Crystal violet staining was assayed as before. Attachment-deficiency was defined as crystal 11

violet staining two standard deviations below that of wild-type. 12

Bacterial Mutagenesis. Targeted deletions were generated in the U112 strain as previously 13

described (9) using the primers in Table A1. Briefly, the regions of the chromosome 5' and 3' to 14

the gene of interest were amplified by PCR. Using splicing by overlap extension (SOE) PCR 15

(44), a kanamycin resistance cassette expressed by the groEL promoter was introduced between 16

these regions of homology. Briefly, ~500bp sequences flanking the targeted gene were 17

amplified and spliced to either end of the gro promoter-resistance cassette construct. The 18

resulting PCR product was transformed into F. novicida strain U112 by chemical transformation 19

and transformants were selected on MMH agar with 30!g/ml kanamycin. Gene deletions were 20

confirmed by sequencing. !secB1, !FTN_1750, and chiA targeted deletion strains were 21

subsequently complemented in cis by re-introducing the wild-type gene into the chromosome at 22

the original locus, along with the CAT cassette chloramphenicol resistance marker, again by 23

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SOE and homologous recombination of a spliced PCR construct. !secB2, !ostA2, !FTN_0308, 1

and chiB deletion mutants were complemented in trans by introducing the wild-type gene, as 2

well as the CAT cassette, into gro-gfp pFNLTP6 (46). ~500bp regions flanking the gfp gene of 3

pFNTLTP were amplified and spliced to the wild-type copy of the gene for complementation 4

with the CAT resistance cassette on the 3’ end. SOE PCR complementation constructs were 5

introduced by homologous recombination with the pFNLTP6 at the NdeI and BamHI sites, 6

removing the gfp gene. The resulting plasmid expressed the complementing gene under the 7

regulation of the constitutive groEL promoter. Complemented strains were selected for growth 8

on 3!g/ml chloramphenicol and also confirmed by sequencing. Complementation plasmids were 9

then chemically transformed into deletion strains. All complementation primers are listed in 10

Table A1. The !chiA!chiB double mutant was constructed using the same method as the single 11

deletion strains, except the chiB gene was replaced with the CAT cassette instead of the 12

kanamycin resistance cassette. 13

RAW264.7 macrophage infections. RAW264.7 macrophages were seeded at 2.5x105 cells per 14

well in 24-well tissue culture plates (Becton Dickinson, Franklin Lakes, NJ) and incubated 15

overnight at 37°C incubation with 5% CO2. Wild-type and mutant U112 strains were grown 16

overnight to stationary phase at 37°C with aeration and diluted to 5x106 colony forming units 17

(CFU) per ml in Dulbecco's Modified Eagle Medium (Gibco, Carlsbad, CA) with 10% fetal 18

bovine serum. For each strain, 1 ml inocula were added to triplicate wells and centrifuged at 730 19

x g for 15 min to mediate attachment. Infected plates were incubated at 37°C with 5% CO2 (time 20

zero) for 0.5h and washed three times with warm media. Three wells per strain were harvested 21

at this time using 0.1% saponin to lyse the cells. CFU were enumerated by serial dilution and 22

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percent recovered was calculated by normalizing to the inocula. Fresh warm media was added to 1

the remaining wells and wells were harvested in triplicate, as above, at 8h and 24h post-infection. 2

Mouse infections. Competitive index (CI) mouse infections were performed as previously 3

described (80) in 6-8 week old female C57BL/6J mice (Jackson Laboratories, Bar Harbor, ME). 4

Mice were infected intradermally or intraperitoneally with equal amounts (5x103 CFU) of wild 5

type and mutant F. novicida in 0.05 ml. Mice were monitored for morbidity and mortality 6

during the course of infection. Mice were sacrificed 2 d post-infection and the spleens were 7

removed and homogenized for CFU enumeration. Competitive indices were calculated as the 8

ratio of mutant to wild type of the output, normalized for the input, and significance was 9

calculated by comparing the CI to 1 (CI of gene with no role in virulence) using one sample t-10

tests. All animal infection experiments were approved by the Institutional Animal Care and Use 11

Committee and the Institutional Biosafety Committee of Stanford University. Deletion mutants 12

for the entire Francisella Pathogenecity Island (FPI) and negative control, hspX chaperone gene 13

were described previously (80). 14

Crab shell attachment. Overnight cultures were grown at 30°C in MMH medium. 15

Approximately 1cm2 pieces of sterile crab shell were inoculated with 2ml of stationary phase 16

cultures in 12-well plates. After 1 h at 30°C, the shells were washed to remove unattached 17

bacteria. Attached bacteria were recovered by vortexing and enumerated for colony forming 18

units (CFUs). All strains were tested in triplicate. Unpaired t-tests were used to determine 19

statistical differences between wild type and mutant counts. 20

Statistical analysis. 21

Statistical analysis was performed using Prism4 software (GraphPad, La Jolla, CA). Unless 22

otherwise stated, unpaired Student’s t tests were applied, and two-tailed P-values are shown. For 23

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mouse CI data, one-sample t-test was used to compare mutant:wild-type bacteria ratio to an 1

expected value of 1. 2

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RESULTS AND DISCUSSION 1

F. novicida forms a biofilm on chitin surfaces. We hypothesized that chitin may be an 2

environmentally relevant surface for the persistence of F. tularensis in nature based on the 3

presence of two well-conserved chitinase genes in the sequenced F. tularensis genomes (Table 4

1). Maintenance of the chiA and chiB genes in F. tularensis subspecies and the related but 5

divergent fish pathogen, Francisella philomiragia, suggested that chitinases provide a selective 6

advantage for Francisella species in nature. F. tularensis subsp. novicida (F. novicida) is a close 7

relative of the highly virulent Type A F. tularensis subsp. tularensis and encodes for both 8

chitinase enzymes. Because F. novicida is genetically tractable, we use this subspecies here as a 9

model to study the molecular aspects of F. tularensis ecology. 10

To test the ability of F. tularensis species to adhere to a chitin-containing surface, we 11

incubated F. novicida with crab shell pieces. Crab shells are rich in chitin, a constituent of 12

various surfaces Francisella species may encounter and subsequently colonize in their natural 13

habitats. These surfaces include copepod and zooplankton shells in fresh water environments 14

and the exoskeletons of arthropod vectors. After 1 h at 30°C, individual and small groups of 15

adhered bacteria were present on the shell surface as visualized by scanning electron microscopy 16

(SEM) (Fig. 1A,B). After one week on the crab shells in the presence of minimal Chamberlain’s 17

defined medium (CDM), without exogenous sugar, three-dimensional bacterial communities 18

were present on the chitin-based surface (Fig. 1C). At higher magnification (Fig. 1D), we saw 19

microcolonies to consist of individual bacteria surrounded by a matrix of extracellular polymeric 20

substance (EPS). The observed community structure suggests F. novicida can attach and 21

proliferate as biofilms on the environmentally relevant surface, chitin. 22

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Although crab shells consist mainly of chitin, they contain additional components, such 1

as other carbohydrates and protein (56). To test if chitin is sufficient to support F. novicida 2

colonization and proliferation, we visualized bacterial attachment and biofilm formation on 3

synthetic chitin films (82). F. novicida attached to lower levels on smooth chitin films compared 4

to the topographically varied crab shells after one hour (Fig. 1A,E). At one week after shift to 5

minimal medium, the surface of the chitin films contained F. novicida microcolonies and EPS 6

extensions (Fig. 1G,H), indicating the initiation biofilm formation. However, the architecture of 7

the bacterial communities on chitin films was not as developed as the communities on the crab 8

shell pieces (Fig. 1A-D), which may be explained by the lower starting population on this 9

surface (Fig. 1C,G). More likely, additional components in the crab shell, like protein, may 10

allow for more rapid expansion of the adhered population. We conclude that chitin is necessary, 11

but not necessarily sufficient for wild-type levels of F. novicida biofilm maturation in the 12

absence of exogenous sugar. 13

F. novicida can utilize GlcNAc as a carbon source for growth. F. novicida persistence and 14

proliferation on chitin surfaces in the absence of exogenous sugar suggested that this pathogen 15

was able to utilize the chitin component of the surface as a nutrient source. To test this, we grew 16

F. novicida in CDM either without added sugar, supplemented with 10mM glucose (a known 17

metabolic substrate for Francisella species), or with 10mM GlcNAc (the monosaccharide end 18

product of chitin hydrolysis) in aerated batch culture. F. novicida growth was negligible in 19

CDM in the absence of an added sugar (doubling time 11.25h). In contrast, F. novicida grew in 20

CDM supplemented with 10mM glucose with a doubling time of 63 min. Similarly, F. novicida 21

grew in CDM supplemented with 10mM GlcNAc (doubling time 76 min.). The high 22

proliferation of F. novicida on chitin surfaces (Fig. 1) may therefore be explained by the ~11-23

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fold increase in growth rate between F. novicida grown in CDM with GlcNAc compared to 1

CDM without sugar. We conclude that F. novicida can metabolize GlcNAc, suggesting that 2

hydrolysis of chitin by chitinases to generate GlcNAc (36) may provide a local nutrient source 3

for persistence and growth. 4

Chitinase genes facilitate F. novicida growth on chitin surfaces. To further address the 5

importance of chitin as a non-host niche for Francisella in nature, we constructed F. novicida 6

mutants lacking either of the chitinase genes, !chiA and !chiB, and a mutant lacking both 7

chitinases. Hager et al. demonstrated that the F. novicida homologs of these enzymes that 8

contain chitin-binding domains are secreted and bind to chitin beads (29). We expected these 9

deletion mutant strains to be attenuated for persistence and biofilm formation on chitin surfaces 10

if F. tularensis species have evolved to form biofilms on chitin surfaces to scavenge carbon. 11

Indeed, the !chiA and !chiB deletion mutants were attenuated for colonization of crab shells 12

when incubated in CDM without sugar. Although the chitinase mutant bacteria attached to chitin 13

to the same extent as wild-type F. novicida at 1 h (data not shown), we recovered 16- and 15-fold 14

fewer !chiA and !chiB mutant bacteria compared to wild-type F. novicida (P<0.001), 15

respectively, after 2 days colonization on crab shells (Fig. 2A). Furthermore, we recovered the 16

same number of !chiA!chiB double chitinase mutant bacteria compared to the !chiA or !chiB 17

single chitinase mutant strains (Fig. 2A), suggesting that the two chitinase genes act in the same 18

metabolic pathway, as predicted by KEGG pathway analysis (34). The abilities of the !chiA and 19

!chiB mutant bacterial strains to grow on chitin was restored by the reintroduction of wild-type 20

copies of each chitinase gene into the coinciding mutant strain as measured by increased crab 21

shell colonization to near wild-type F. novicida levels (Fig. 2A). The ability of the chitinase 22

mutant strains to persist at low levels could be due to the utilization of the amino acids present in 23

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the CDM medium. Alternatively, natural degradation of the crab shell during the experiment 1

could liberate enough free GlcNAc to enable the bacteria to persist, but not replicate. 2

Regardless, the highly significant difference between wild-type and chitinase mutant bacteria 3

suggests that chitinase activity strongly contributes to F. novicida persistence on chitin in 4

otherwise carbon-limiting conditions. 5

We postulated that the inability of chitinase mutant bacteria to convert chitin to the 6

useable metabolite, GlcNAc, explains their attenuated colonization on chitin. Indeed, the 7

inability of the chitinase mutants to colonize crab shells was alleviated by the addition of 10mM 8

GlcNAc to the exogenous medium (Fig. 2B), indicating that the chitinase mutant bacteria 9

possess the determinants required to colonize a chitin surface, but lack the ability to generate a 10

useable carbon source in order to proliferate. The 13-fold decrease in recovered wild-type F. 11

novicida when GlcNAc was added (Fig. 2) is consistent with microarray data published for V. 12

cholerae demonstrating that when this pathogen was grown in the presence of excess GlcNAc, 13

the pili and chitinases required to colonize this surface were repressed (49). 14

We next compared the architecture of the communities formed by the chitinase mutants 15

on crab shells or chitin films in the absence of exogenous sugar for 1 week by scanning electron 16

microscopy. In contrast to wild-type F. novicida, the chitinase mutants were present as single 17

bacteria or small, mostly monolayer, clusters of bacteria (Fig. 3). We conclude that F. novicida 18

biofilm formation on chitin in the absence of exogenous sugar requires functional chitinase 19

enzymes. Unlike motile V. cholerae that can chemotax towards nutrients, F. tularensis species 20

are non-flagellated and non-motile under laboratory conditions (14). Therefore, the ability of 21

Francisella species to adhere and colonize chitin may represent a single mechanism for survival 22

in nutrient poor non-host environments. Growth on chitin may trigger a specific biofilm 23

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program of genes that promote the retention of scavenged GlcNAc in the local 1

microenvironment for use by F. tularensis. 2

Beyond scavenging carbon in the environment, the secreted chitinases that are vital for 3

biofilm formation on chitin could be important for the establishment of the arthropod infection, 4

similar to the malaria parasite Plasmodium falciparum (79). The P. falciparum chitinase allows 5

the parasite to penetrate the chitin-containing peritrophic matrix surrounding the blood meal in 6

the mosquito midgut and establish the infection. Efforts to target this chitinase to block 7

transmission of malaria are ongoing (69, 73). We are currently working to discern the role(s) of 8

F. tularensis chitinases in arthropod vectors. 9

Characterization of biofilm development by Francisella species. F. tularensis chitin 10

utilization provides insight into potential persistence mechanisms of this highly virulent 11

pathogen. The missing piece to our model was the proteins that promote attachment to chitin 12

surfaces. We established in vitro systems for studying F. tularensis biofilm formation to aid in 13

identifying attachment determinants. In vitro biofilms on abiotic surfaces provided a model 14

system to characterize and genetically dissect F. novicida biofilm formation and test the ability 15

of other pathogenic F. tularensis strains to similarly attach and proliferate on a surface. 16

We incubated GFP-labeled F. novicida in the flow cell system (12) to confirm in vitro 17

formation of these bacterial communities under flow conditions. Bacterial attachment and 18

surface growth at ambient temperature (20-22°C) and a flow rate of 0.1ml/min was analysed by 19

confocal laser scanning microscopy (CLSM) at various timepoints (24h, 48h, 72h, 96h and 120h) 20

(Fig. 4). We observed the formation of a matt-like biofilm with an average depth of 21

approximately 15µm. This architecture of flow cell grown F. novicida biofilms was similar to 22

that reported for other Gram-negative species, including the related "-proteobacterium 23

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Shewanella oneidensis (76) and !-proteobacterium Caulobacter crescentus (20). Our results 1

indicate that F. novicida is able to form biofilms on an abiotic surface, such as glass, with similar 2

architecture to that observed on chitin (Fig. 1). These results are consistent with the report by 3

Hassett et al. indicating that LVS can form biofilms on glass coverslips in the absence of flowing 4

media (30). 5

A modified O’Toole and Kolter microtiter assay (59) was utilized to establish a high 6

throughput model for F. tularensis biofilm formation. This assay measures adhered biomass 7

under static conditions by crystal violet stain. F. novicida and F. tularensis subsp. holarctica 8

live-vaccine (LVS) strains were grown at 26°C and 37°C in 96-well microtiter plates. Recent 9

work by Horzempa et al. found that the LVS strain demonstrated different expression profiles at 10

these two temperatures (31). The OD570 (Fig. 5A,B) and crystal violet staining (CV570), (Fig. 11

5C,D) were measured over 152 h. Both F. novicida and LVS strains showed increased crystal 12

violet staining over time when grown at 26°C and 37°C, indicating increased accumulation of 13

adhered biomass. This result is consistent with our finding that F. novicida forms biofilms when 14

adherent to an abiotic surface (Fig 4). At both temperatures assayed, we observed a decrease in 15

crystal violet staining (Fig. 5C,D) concurrent with F. novicida and LVS entering stationary phase 16

(Fig. 5A,B). This result suggested that the biofilms were undergoing dispersion (75), a process 17

of biofilm dissolution and re-seeding occurring during decreased oxygen tension and nutrient 18

deprivation. Similar dispersal did not occur in the flow cell system grown F. novicida biofilms 19

(Fig. 4), presumably because the population was constantly provided an undepleted carbon and 20

oxygen source under flow conditions. 21

Type A Francisella strains form biofilms in the microtiter plate assay. A high percentage of 22

tularemia morbidity and mortality is caused by infection with F. tularensis subsp. tularensis 23

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(Type A) strains (22). These strains have a very low infectious dose, and as few as ten 1

organisms can cause a lethal infection in humans (19). Molecular subtyping of Type A strains 2

has identified two distinct subtypes (A1 and A2) with specific geographic distributions (40). 3

Type A1 strains are primarily found in the Eastern United States, while Type A2 strains are 4

almost exclusively isolated in the West. The O’Toole and Kolter assay demonstrated that these 5

highly virulent strains were able to form biofilms to similar levels as F. novicida and LVS strains 6

(Fig. 6). SchuS4 (Type A1) and FT-10 (Type A2) F. tularensis subsp. tularensis strains reached 7

similar optical densities as LVS (Type B) when grown under static conditions (Fig. 6), while 8

SchuS4 and FT-10 exhibited higher crystal violet staining at 24 h (P<0.05), implying increased 9

biofilm formation of Type A strains (Fig. 6). F. novicida CV600 staining was approximately two-10

fold higher than the other strains tested (P<0.001). However, the optical density of the F. 11

novicida culture was 2.5-fold higher than the other strains at 24h. Similar crystal violet staining 12

by Type A1 and Type A2 strains compared to the Type B LVS strain suggests that biofilm 13

formation may be pertinent to the survival of pathogenic F. tularensis strains in the environment. 14

Screen for biofilm-deficient mutants identifies novel genes important for F. novicida 15

biofilm formation. We screened a two-allele transposon library (BEI Resources, Manassas, 16

VA) that represented two or more transposon-insertion mutants per non-essential gene in the F. 17

novicida genome to elucidate the genetic determinants of F. novicida interaction with abiotic and 18

biotic surfaces. To facilitate high-throughput screening, individual insertion mutants were 19

assayed for biofilm formation in the microtiter assay established above rather than on chitin. We 20

defined biofilm-deficient mutants as strains where crystal violet staining was two standard 21

deviations below the mean of the plate. We eliminated mutant strains that exhibited a significant 22

growth defect from further characterization. In total, we identified 98 F. novicida transposon-23

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insertion mutants, representing 88 genes that were attenuated for biofilm formation (Table A2). 1

To elucidate pathways important for F. novicida surface attachment and growth, we assigned 2

gene ontology classifications to the genes identified in the biofilm screen 3

(www.biohealthbase.org) (Fig. A1). Roles for the 64 annotated genes included protein secretion, 4

various metabolic pathways, signal transduction, protein transport, and cell envelope biogenesis. 5

Although many Gram-negative and Gram-positive bacteria can form biofilms, the bacterial 6

mechanisms utilized to facilitate these communities vary (4, 45, 55, 59, 78). For example, the 7

role of Type-IV pili and flagella in biofilm formation is well documented (58, 65). However, 8

little is known about attachment and surface growth during biofilm maturation of non-motile 9

bacteria. By characterizing the roles of the genes we identified in this study, including the 10

approximately 25% with no annotated function, we aim to elucidate alternate methods of 11

environmental persistence by non-motile bacteria. 12

Sec-dependent secretion functions in initial attachment during F. novicida biofilm 13

formation on abiotic and biotic surfaces. We were particularly interested in the four 14

transposon-insertion mutants we identified in the Sec translocon complex involved in protein 15

export from the cytoplasm (25). The core components of the Sec translocon in Escherichia coli 16

are the SecYEG protein channel and the SecA ATPase motor protein (10). Due to the 17

pleiotropic roles of general protein secretion in bacteria, components in this apparatus are 18

considered essential in other Gram-negative organisms (24, 43). The Sec translocon in F. 19

novicida is comprised of 13 proteins, but only the four genes we identified in our screen were 20

represented in the two-allele library; the secA motor ATPase and secG pore genes, as well as the 21

secB1 and secB2 genes that encode for chaperones which specifically target pre-proteins to SecA 22

(77). Additionally, we identified 18 transposon biofilm mutant clones, representing 14 genes 23

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that are predicted to encode for proteins with secretion signals based on the signal sequence 1

detection algorithm, SignalP (7) (Table 2). 2

We hypothesized from the results of our genetic biofilm screen and a secondary 3

attachment assay that proteins secreted by the Sec translocon may represent novel mediators of 4

F. novicida adhesion, a process that has not been characterized. We confirmed that transposon 5

mutants in the secretion apparatus were deficient for biofilm formation (Fig. 7A) and attachment 6

(Fig. 7B). Deletion mutants in secB1 and secB2 were constructed while deletions in secA and 7

secG could not be generated; suggesting that these genes are essential and the transposon-8

insertion mutations present in the library represent an incomplete loss of gene function. 9

Additionally, attempts to construct a double deletion of secB1 and secB2 did not yield viable 10

colonies. Growth curves performed with the !secB1 and !secB2 mutants showed no growth 11

defect in batch culture compared to wild-type F. novicida and microscopic analysis of cell 12

morphology revealed no alternations in bacterial shape (data not shown). Both the !secB1 and 13

!secB2 mutants were deficient in biofilm formation (Fig 7C) and attachment (Fig 7D) when 14

grown in MMH. The !secB1 and !secB2 mutant phenotypes were restored to wild-type 15

attachment and biofilm formation levels when wild-type copies of secB1 and secB2 were added 16

back to the deletion mutants (Fig. 7C,D). These experiments were also performed in CDM to 17

confirm that the role of Sec-dependent secreted factors in biofilm formation was not limited to 18

growth in a nutrient-rich environment (Fig. 7E,F). Our data indicate that Sec-dependent 19

secretion is important for F. novicida attachment to abiotic surfaces and biofilm formation. We 20

therefore postulated that Sec-secreted proteins represent novel mediators of F. novicida 21

adherence. 22

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The 18 Sec-dependent transposon-insertion mutants (Table 2) were all defective for 1

biofilm formation (Fig. 7A) and initial attachment (Fig. 7B) based on crystal violet staining, 2

confirming our screen results. Surprisingly Type-IV pili genes, known mediators of biofilm 3

formation in Gram negative bacteria, were not among the Sec-secreted factors identified and 4

were found to be dispensible for F. novicida biofilm formation upon further study (data not 5

shown). 6

We focused on four of the secreted factors with homologs in all F. tularensis subspecies 7

and were highly attenuated for biofilm formation when deleted; FTN_0308, FTN_0713, 8

FTN_0714 and FTN_1750. FTN_0713 (ostA2), FTN_0714 and FTN_1750 were all identified at 9

least twice in the biofilm screen. We selected FTN_0308 due to the strong biofilm phenotype of 10

the one transposon-insertion mutant that was identified in the genetic screen (Fig. 7A,B). We 11

constructed deletion mutants in each of these genes and tested for attachment and biofilm 12

formation. All four mutants were defective in initial attachment and biofilm formation in both 13

rich and defined media (Fig 7C-F). The !ostA2, !FTN_1750 and !FTN_0308 mutants were 14

complemented for attachment and biofilm attenuation by re-introduction of the deleted genes in 15

cis into the chromosome or in trans by expressing the gene in pFNLTP6 using the constitutive 16

gro promoter. The !FTN_0714 mutant could not be complemented for technical reasons, likely 17

due to the length of the complementation PCR product (~8kb). Taken together, our data indicate 18

that initial attachment during Francisella biofilm formation is facilitated by proteins secreted by 19

the Sec-dependent secretion system. 20

The protein encoded by FTN_0713 (ostA2) has significant homology (E-value 6e-64) to 21

organic solvent tolerance proteins involved in lipopolysaccharide (LPS) modification (8). 22

Although ostA2 homologs have not been implicated in biofilm formation, LPS chemistry has 23

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been shown to influence attachment during biofilm formation in other bacteria (4, 16, 23). The 1

unique structure of Francisella species LPS (28) could contribute to adhesion of F. tularensis to 2

non-mammalian surfaces. FTN_1750 is a putative acyltransferase with strong homology (E-3

value 4e-27) to acylhomoserine-lactone biosynthesis enzyme, HdtS, suggesting that this protein 4

may function in quorum sensing, a cell-cell communications process that regulates biofilm 5

formation under certain conditions (38). 6

While F. novicida biofilm genes were identified by screening for mutants defective for 7

adherence and biofilm formation on polystyrene, we identified two novel putative chitin-binding 8

proteins, FTN_0308 and FTN_0714. The protein encoded by FTN_0714 is annotated as a 9

hypothetical lipoprotein (BioHealthBase). The SMART domain prediction algorithm (42, 68) 10

indicates that FTN_0714 contains repeating polycystic kidney disease-family domains conserved 11

from archae through mammals that facilitate adhesion (33). This domain-family plays a role in 12

the binding and hydrolysis of chitin by the chiA chitinase of aquatic bacterial strain Alteromonas 13

0-7 (60). FTN_0308 is annotated to encode for a hypothetical protein with unknown function 14

(www.biohealthbase.org). However, the Phyre protein-folding prediction algorithm (35) 15

indicates a structural homology to the Streptomyces chitinase C chitin-binding domain and the C-16

terminus contains homology to F17c-family bacterial adhesins. We are currently determining 17

the specific roles that these two gene products may have in attachment to both abiotic and chitin 18

surfaces. 19

F. tularensis species genomes contain an annotated chitin-binding protein, cbpA, that was 20

not identified by our biofilm screen. This gene product may specifically mediate association 21

with chitin. Additionally, we did not identify chiA and chiB in our screen despite their conserved 22

Sec-dependent secretion signals and role in biofilm formation on chitin. We would not expect 23

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chitinases to mediate biofilm formation on polystyrene, however. This result was confirmed 1

using clean deletion mutant stains (data not shown). 2

Sec secretion mutants are not attenuated in murine models of infection. While no evidence 3

of F. tularensis biofilm formation inside of mammalian hosts exists, bacteria often utilize 4

proteins to attach to both environmental and host surfaces (65, 67, 70). Attachment of F. 5

tularensis in any context is poorly understood. We, therefore, tested if the biofilm attachment 6

factors we describe also mediate host tissue association using in vitro and in vivo infection 7

models. F. tularensis species are primarily found within macrophages in a mammalian host (27). 8

Therefore, RAW264.7 macrophage-like cells were infected at a multiplicity of infection of 20:1 9

with wild-type F. novicida or the Sec-dependent secretion mutants. At 0.5 h post-infection, non-10

cell associated F. novicida were washed away and the remaining bacteria were recovered and 11

enumerated. No statistical differences in CFU counts were observed (Fig. 8A), suggesting that 12

the mutants that are defective for attachment to polystyrene and chitin were still able to 13

efficiently associate with eukaryotic cells. Intracellular replication was monitored in the 14

presence of extracellular gentamicin for 8- and 24h (Fig. 8B). Wild-type F. novicida and all 15

mutants showed approximately 100–fold replication at 24h compared to the initial 0.5 h counts. 16

Thus, the mutants successfully entered and replicated within macrophages, demonstrating that 17

the Sec secretion biofilm mutants that we characterized are not deficient for attachment to, or 18

replication within macrophages. 19

To test the potential role of secB1, secB2, ostA2, FTN_0308, FTN_0714 and FTN_1750 20

during a systemic mouse infection, we infected C57BL/6J mice with a 1:1 mixture of 5x103 21

colony forming units of wild-type and deletion mutant bacteria. Competitive indices (CI’s) for 22

each wild-type/mutant combination were obtained for both intradermal (ID) and intraperitoneal 23

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(IP) routes of infection. Mutants that are not attenuated in mice should have a CI of one, i.e., 1

equal numbers of wild-type and mutant bacteria are recovered in the tissue at the time of harvest, 2

as is observed for the previously described !hspX strain (80). As a positive control, we included 3

a F. novicida !FPI mutant that lacks the entire Francisella pathogenecity island (80). As 4

expected, this mutant was severely attenuated in mice (Fig. 8C,D). However, none of the Sec 5

secretion biofilm mutants demonstrated a CI value statistically different from one via either route 6

of infection (Fig. 8C,D). Additionally, no defect was observed in the spread of Sec secretion 7

mutants to systemic tissues such as the liver and spleen after ID inoculation (data not shown). 8

Our data indicate that these genes that are crucial for association to non-mammalian surfaces do 9

not contribute to local or systemic colonization of mammalian hosts. FTN_0713 (ostA2), the 10

putative LPS-modification gene, was identified by Kraemer et al. in a negative selection screen 11

for F. novicida mutants attenuated for infection via intranasal inoculation of mice, indicating that 12

this mutant may be more sensitive to the innate immune response in the lung, (e.g., antimicrobial 13

peptides) due to an altered LPS (39, 80). Transposon-mutants for secA and secE were identified 14

by Su et al. to be involved in lung colonization (72). These attenuated phenotypes for the non-15

redundant Sec translocon genes imply that Sec secreted proteins other than those characterized 16

here do influence host colonization. The lack of attenuation for the deletion mutants in secB1 17

and secB2 in the virulence assays tested here supports the idea that these two genes encode for 18

redundant function. 19

F. novicida biofilm determinants also play a role in attachment to chitin-based surfaces. 20

Given Francisella species induce biofilm formation on both abiotic and chitin surfaces, we 21

hypothesized that the attachment determinants we identified for association with polystyrene 22

may also facilitate attachment to chitin. After a 1 h incubation at 30°C, an average of 3.33 x 107 23

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CFU/ml wild-type F. novicida were attached to the crab shell pieces (Fig. 9). Sec secretion 1

mutants were 5.6- to 16.2-fold attenuated for attachment to this chitin-based surface compared to 2

wild-type bacteria (P<0.01), confirming that Sec-secreted proteins contribute to attachment to 3

chitin-based surfaces. The specificity of these adherence factors for non-mammalian surfaces 4

further supports our suggestion that F. tularensis biofilm formation in nature has evolved to 5

promote this pathogen’s survival outside of a host; potentially by facilitating chitin utilization. 6

From our collective data we propose a model in which these early determinants of 7

biofilm formation allow for association with chitin surfaces in nature. Through this interaction, 8

F. tularensis chitinases have access to this substrate and provide bacteria with GlcNAc, which is 9

utilized for growth in nutrient-limiting environments. Biofilm maturation on chitin would then 10

create a local microenvironment enriched for this carbon source, providing a non-host niche for 11

this zoonotic pathogen. 12

The ability of F. tularensis to form biofilms on chitin may also provide the bacterium 13

resistance to grazing by fresh water protozoa. For chitin-colonizer V. cholerae, biofilm 14

formation was shown to reduce grazing by flagellate organisms compared to planktonic bacteria 15

(48). Thelaus et al. found that F. tularensis subsp. holarctica had increased resistance to both 16

ciliate and flagellate protozoa compared to E. coli (74). Although the role of biofilm formation 17

on predation was not addressed, this observation suggests that F. tularensis may actively prevent 18

protozoal grazing in nature. Coupled with the ability to survive in nutrient-limited aquatic 19

environments, biofilm-mediated resistance to predation could contribute to F. tularensis 20

persistence in the environment and allow for prolonged transmission of this pathogen. 21

We provide here the first extensive characterization of F. tularensis biofilm formation 22

and explore how these communities may promote environmental persistence and transmission on 23

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chitin surfaces. The F. novicida biofilm genes we describe contribute to the ability of this 1

pathogen to colonize a surface it may encounter in nature. Very little is known about how and 2

where Francisella species persist in nature when not replicating within a host. Our findings may 3

help explain how tularemia outbreaks that have been attributed to fresh water crustaceans (2, 17) 4

occur. Additionally, chitin utilization may support F. tularensis persistence on other arthropods 5

such as zooplankton, copepods, and biting arthropod vectors. A study of F. tularensis survival in 6

artificial water found that the presence of chitinous fresh water shrimps, mullosks, diatoms, or 7

zooplankton promoted sustained viability of this pathogen for an additional week to one month 8

in nutrient-poor water (52). Survival on environmental chitin may therefore serve as a reservoir 9

for disease transmission during seasonal tularemia outbreaks. Palo et al. identified a strong 10

epidemiological correlation between areas with low water turnover and human cases of tularemia 11

(64). These researchers postulated low water turnover as an environmental cue for a burst of F. 12

tularensis replication. As with cholera outbreaks (21), conditions that promote interaction of F. 13

tularensis with chitin surfaces on which the bacteria can replicate may seed infection. Further 14

study of F. tularensis biofilm formation and the role of these communities in chitin colonization 15

could clarify the open question of the location of the F. tularensis environmental reservoir. 16

17

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ACKNOWLEDGEMENTS 1

We thank Melanie Blokesch for her generous gifts of reagents and technical assistance, Gary K. 2

Schoolnik for his thoughtful discussions and for providing the chitin films. Jonathan W. Jones 3

and Thomas Henry for help with mouse experiments and Carmen D. Cordova for assistance with 4

the biofilm CLSM imaging. Jean Celli graciously provided the SchuS4 strain. J.J.M was 5

supported by National Science Foundation and Department of Homeland Security graduate 6

fellowships, as well as a National Institutes of Health Cell and Molecular Biology training grant. 7

8

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FIGURE LEGENDS 1

Figure 1. F. novicida biofilm formation on chitin surfaces. Images display SEM visualization 2

of F. novicida colonization of crab shell pieces (A-D) and synthetic chitin films (E-H). 3

Individual attached bacteria and small attached microcolonies were observed on the crab shell 4

pieces at one hour (A,B). After one week, typical 3D biofilm architecture was observed, 5

consisting of bacteria surrounded by an EPS matrix (C,D). Similar results were obtained after 6

one hour (E,F) and one week (G,H) on synthetic chitin. Scale bar is 20µm for lower 7

magnification images (left column) and 5µm for higher magnification images (right column). 8

9

Figure 2. Chitinase mutants are attenuated for chitin colonization in the absence of 10

exogenous sugar. Stationary phase wild-type and chitinase mutant bacteria were allowed to 11

adhere for 1 h to crab shell pieces. Equivalently adhered strains were allowed to colonize these 12

chitin surfaces in CDM with or without GlcNAc at 30°C. Triplicate samples were harvested 2 d 13

post-inoculation and enumerated for CFU. Chitinase mutant F. novicida (white) were recovered 14

at statistically lower levels than wild-type bacteria (black), (P<0.001) when incubated in CDM 15

(A), but in equivalent numbers in CDM with GlcNAc (B). Addition of wild-type chiA and chiB 16

genes to deletion mutant strains (grey) complemented the chitin colonization defects observed 17

during colonization in CDM without GlcNAc (A). 18

19

Figure 3. Chitinase genes are required for biofilm architecture on chitin surfaces during 20

nutrient stress. Images show representative colonization by wild-type and chitinase mutant 21

strains on crab shells (A-D) or synthetic chitin films (E-H). Bacteria were allowed to attach for 1 22

h and then incubated for one week at 30°C before being processed for SEM. In contrast to 23

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extensive 3D biofilm development in wild-type F. novicida, the chitinase mutants were present 1

as single bacteria or small clusters of bacteria on both natural and synthetic chitin. Scale bar is 2

10µm. 3

4

Figure 4. Francisella forms a matt-like biofilm under flow conditions. GFP expressing F. 5

novicida grown at room temperature (20-22°C) were imaged daily in flow cells run at 0.1ml/min 6

using confocal laser scanning microscopy. Representative images from triplicate experiments 7

are shown. At 24h (A), small groups of bacteria are present. Over the next 48h (B,C), a uniform 8

monolayer of bacteria is observed on the surface. By 96h (D), depth in the biofilm is observed 9

and at 120h (E) the biofilm reached an average thickness of 15µm. Scale bar is 15.2µm. 10

11

Figure 5. Kinetics of F. tularensis biofilm formation under static conditions. A modified 12

O’Toole and Kolter assay was performed to compare the kinetics and relative levels of biofilm 13

formation for F. novicida (solid circles) and LVS (open circles). Bacterial growth (A,B) and 14

crystal violet staining (C,D) were determined over time at 26°C (A,C) and 37°C (B,D) by OD570 15

readings. Both F. tularensis strains were found to acquire crystal violet stain at both 16

temperatures. Growth and crystal violet staining were faster at 37°C for both strains. 17

18

Figure 6. Biofilm formation by virulent F. tularensis subspecies tularensis strains. F. 19

novicida, LVS and Type A strains SchuS4 and FT-10 were assayed for growth and crystal violet 20

staining at 24h post-inoculation. Culture OD600 (A) and crystal violet staining (B) were 21

determined after static growth at 37°C. F. novicida demonstrated increased growth kinetics and 22

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crystal violet staining compared to the other strains (P>0.001). Virulent SchuS4 and FT-10 1

strains exhibited significantly higher crystal violet staining compared to LVS. 2

3

Figure 7. Sec-secreted factors mediate initial attachment during biofilm formation. 5 4

transposon-insertions representing mutants in 4 genes in the Sec translocon (grey) and 18 5

transposon-insertions representing mutants in 14 genes in putative secreted factors (white) 6

identified in the forward genetic screen were tested in triplicate compared to wild-type F. 7

novicida (black) for 8h biomass accumulation (A) and 1 h initial attachment (B). Multiple 8

transposon-mutants were tested for genes identified more than once in the screen. Adherence of 9

biomass at 8h was used a measurement for biofilm formation. Attachment was assessed by 10

crystal violet stain 1 h post-inoculation of stationary phase cultures. Targeted mutants in 11

selected representative genes (white) showed similar defects in biofilm formation (C, E) and 12

attachment (D, F) compared to wild-type F. novicida (black) when grown in MMH and CDM, 13

respectively, based on crystal violet staining. Complementation of deleted genes (grey) restored 14

mutants to wild-type levels in all cases. Bars represent the mean and the lines indicate standard 15

deviation calculated from triplicate samples of a representative experiment. Each experiment 16

was repeated in triplicate. No data (ND) was obtained for FTN_0714 complementation due to 17

technical difficulties. 18

19

Figure 8. The Sec translocon and secreted factors do not influence F. novicida virulence. 20

Sec secretion targeted deletion mutants were assessed in in vitro and in vivo models for F. 21

tularensis virulence. Entry efficiency of F. novicida strains into RAW264.7 macrophage-like 22

cells was measured as the percent of inocula recovered from inside the cells 30min post-infection 23

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(A). Intracellular replication of wild-type and mutant bacteria was assessed as fold-replication 1

compared to 30 min counts at 8 h and 24 h post-infection (B). The ability of mutants to colonize 2

the skin after intradermal (C) and the spleen after intraperitoneal (D) routes of inoculation was 3

determined by competitive indices in C57BL/6J mice 2 d post-infection. For all virulence 4

assays, no difference was observed between the Sec secretion biofilm mutants and wild-type F. 5

novicida. 6

7

Figure 9. Biofilm mutants are attenuated for attachment to chitin-based crab shell pieces. 8

Stationary phase cultures of secB1, secB2, ostA2, FTN_0308, FTN_0714, and FTN_1750 9

deletion mutants were allowed to attach for 1 h to sterile crab shell pieces. Attached bacteria 10

were enumerated for CFU in triplicate samples. Sec secretion biofilm mutants were found to 11

attach statistically lower (P>0.01) than wild-type F. novicida by unpaired t-test. 12

13

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Table 1 – Francisella species chitinase genes

Strain chiA homolog (E-value)* chiB homolog (E-value)

*

F. tularensis subsp. tularensis SchuS4 FTT0715 (9e-66) FTT_1768c (2e-15)

F. tularensis subsp. tularensis FSC198 FTF0715 (3e-70) FTF_1768c (2e-15)

F. tularensis subsp. holarctica LVS FTL_1521 (9e-66) FTL_0093 (1e-15)

F. tularensis subsp. holarctica OSU18 FTH_1471 (2e-68) FTH_0088 (1e-15)

F. tularensis subsp. novicida U112 FTN_0627 (4e-69) FTN_1744 (4e-14)

Francisella philomiragia Fphi_0215 (1e-66) Fphi_0864 (1e-15) * E-value based on comparison to glycosyl hydrolase 18 family chitinases

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Table 2- Sec translocon and Sec-dependent secreted proteins involved in biofilm formation

FTN Well ID

a Gene Gene Product Biological Process Sec Secretion

b

FTN_0090 4F07 acpA acid phosphatase fatty acids and lipids

metabolism

Secreted

FTN_0100 20C12 hypothetical membrane protein

hypothetical - novel Secreted

FTN_0109 14G06 protein of unknown function unknown function - novel Secreted

FTN_0121 26G09 secB1 preprotein translocase, subunit B

motility, attachment and secretion structure

Translocon

FTN_0121 4F06 secB1 preprotein translocase, subunit B


Translocon

FTN_0191 19 E06 polar amino acid uptake transporter

transport - amino-acid Secreted

FTN_0304 20C11 pilus assembly protein motility, attachment and secretion structure

Secreted

FTN_0308 19H06 membrane protein of unknown function

unknown function - novel Secreted

FTN_0357 21B08 pal peptidoglycan-associated lipoprotein, OmpA family

transport - drugs / antibacterial compounds

Secreted

FTN_0429 14G12 conserved protein of unknown function

unknown function - conserved Secreted

FTN_0635 25C04 serine-type D-Ala-D-Ala carboxypeptidase

cell wall / LPS / capsule Secreted

FTN_0672 12G03 secA preprotein translocase, subunit A (ATPase, RNA

helicase)


Translocon

FTN_0713 14C04 ostA2 organic solvent tolerance

protein OstA


FTN_0713 21 h10 ostA2 organic solvent tolerance protein OstA


FTN_0713 26 E07 ostA2 organic solvent tolerance protein OstA


FTN_0714 12G01 protein of unknown function unknown function - novel Secreted

FTN_0714 27C09 protein of unknown function unknown function - novel Secreted

FTN_1093 18A05 protein of unknown function unknown function - novel Secreted



FTN_1510 1 E01 secB2 preprotein translocase, subunit B


Translocon

FTN_1630 13C11 secG preprotein translocase, subunit G, membrane

protein


Translocon

FTN_1750 19H02 acyltransferase fatty acids and lipids metabolism

Secreted

FTN_1750 23D04 acyltransferase fatty acids and lipids metabolism

Secreted

a Well ID annotation from BEI Resources F. novicida Two-Allele Transposon Library. b Genes labeled translocon are structural components of Sec-dependent secretion. Secreted proteins were predicted using SignalP algorithm.

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CDM ( E ) ( F )N DW T s e c B 1 s e c B 2 o s t A 2F T N _0 3 0 8F T N _0 7 1 4F T N _1 7 5 00 . 00 . 10 . 20 . 30 . 40 . 5CV570W T s e c A s e c B 1 s e c B 1 s e c B 2 s e c G a c p A o s t A 2 o s t A 2 o s t A 2 p a lFT N _0 1 0 0FT N _0 1 0 9FT N _0 1 9 1FT N _0 3 0 4FT N _0 3 0 8FT N _0 4 2 9FT N _0 7 1 4FT N _0 7 1 4FT N _1 0 9 3FT N _1 4 7 6FT N _1 5 0 3FT N _1 7 5 0FT N _1 7 5 00 . 00 . 20 . 40 . 60 . 8CV570T ransposonM ut ant s

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